Preparation of chaperone-loaded neural stem cell-derived extracellular vesicles to reduce protein aggregation in Huntington’s disease cellular models

Summary Here, we present a protocol using genetic engineering techniques to prepare small extracellular vesicles (sEVs) enriched in the chaperone protein DNAJB6. We describe steps to prepare cell lines overexpressing DNAJB6, followed by the isolation and characterization of sEVs from cell conditioned media. Further, we describe assays to examine effects of DNAJB6-loaded sEVs on protein aggregation in Huntington’s disease cellular models. The protocol can be readily repurposed to study protein aggregation in other neurodegenerative disorders or extended to other therapeutic proteins. For complete details on the use and execution of this protocol, please refer to Joshi et al. (2021).1


SUMMARY
Here, we present a protocol using genetic engineering techniques to prepare small extracellular vesicles (sEVs) enriched in the chaperone protein DNAJB6. We describe steps to prepare cell lines overexpressing DNAJB6, followed by the isolation and characterization of sEVs from cell conditioned media. Further, we describe assays to examine effects of DNAJB6-loaded sEVs on protein aggregation in Huntington's disease cellular models. The protocol can be readily repurposed to study protein aggregation in other neurodegenerative disorders or extended to other therapeutic proteins. For complete details on the use and execution of this protocol, please refer to . 1

BEFORE YOU BEGIN
This protocol describes methods for isolating small extracellular vesicles (sEVs) from neural stem cell line C17.2 overexpressing the chaperone protein DNAJB6b and studying the effect of these sEVs on protein aggregation in in vitro cellular models of Huntington's disease.

Timing: 4 days
This section describes how to prepare plasmids expressing XPackTM-tagged DNAJB6.
1. Purchase XPackä CMV-XP-MCS-EF1-Puro Cloning Lentivector from SBI biosciences (XPAK510PA-1). For more details about the vector, please refer to the company manual (https://www.systembio.com/wp/wp-content/uploads/2020/10/Manual_XPack_WEB-1.pdf). 2. Amplify GFP and GFP-DNAJB6 fragments by PCR using pcDNA5/FRT/TO GFP-DNAJB6b (Addg-ene#19501). Follow the recipe below for PCR-3. Digest the respective PCR segments and the parent lentiviral vector with XhoI-HF and EcoRI-HF. The PCR fragments are 733 bp (GFP) and 1,471 bp (GFP-DNAJB6) in size. Prepare the digestion reaction mixture as shown below and incubate at 37 C for 2 h. 4. Purify the digested PCR products using QIAquick PCR & Gel Cleanup Kit (https://www.qiagen. com/br/resources/download.aspx?id=a72e2c07-7816-436f-b920-98a0ede5159a&lang=en). 5. Run the digested vector on an agarose gel at 200 V until resolved properly to separate the DNA fragment generated by XhoI and EcoRI restriction from the vector. There should be two bands on the gel of differing sizes. 6. Under UV illumination, excise out the bigger band, which represents the remaining vector DNA to be inserted with the PCR amplicons. Using QIAquick PCR & Gel Cleanup Kit (https://www.qiagen. com/br/resources/download.aspx?id=a72e2c07-7816-436f-b920-98a0ede5159a&lang=en), elute the DNA from the gel piece. 7. Insert the GFP and GFP-DNAJB6 PCR fragments into the cut vector using ligation to produce pCMV-XP-GFP-EF1-Puro and pCMV-XP-GFP-DNAJB6-EF1-Puro, respectively. Set up the ligation reaction mixture given below and incubate at RT for 10 min. 8. Transform the ligation mixture into bacteria (DH5a) and plate on agar plates containing ampicillin, incubate for 18 h at 37 C. 9. The next day, screen 10 colonies with a colony PCR using Green GoTaq PCR reaction (Promega) 10. Select three clones that give a clean PCR band at expected size. 11. Inoculate the three clones separately in 3 mL LB medium containing ampicillin and grow overnight at 37 C. 12. Isolate plasmids using Promega plasmid miniprep kit (https://nld.promega.com/resources/ protocols/technical-bulletins/101/pureyield-plasmid-miniprep-system-protocol/). 13. Send clones for DNA sequencing and select one that shows no mutations in the insert sequence. 14. The final vectors are shown in Figure 1. After overnight incubation the cells should be 60% confluent. Wash twice with HBSS and proceed to adding the EV-depleted medium. For T162 flask, add 20 mL medium (which will just cover the surface) to concentrate the sEV sample. CRITICAL: C17.2 neural stem cells multiply very fast. When crowded, they tend to differentiate into elongated neuronal cells. It is important to control the cell density while culturing them for regular passaging as well as extracellular vesicles collection to prevent the presence of differentiated cells. 60% cell confluency is a good moment for the addition of EV-depleted medium, and 2 days incubation time is sufficient to get a good amount of EVs from 10 T162 flasks (2 mg/mL, 50 mL total volume).

OPEN ACCESS
We present here a method of monoclonal cell preparation using a paper disc method, which is a labor-and cost-effective method of generating a monoclonal cell line without the need for sophisticated equipment such as a FACS sorter. CRITICAL: In our hands, lipofection and lentiviral transduction did not result in stable C17.2 cell lines with high exogenous protein expression (which is necessary for high protein loading in EVs, which in turn would improve the therapeutic efficacy of EVs). Nucleofection gave the best results in terms of high protein expression. Hence, we have used nucleofection in this protocol.
Note: Nucleofection, combining cell type-specific Nucleofector Solution and electrical pulses, facilitates DNA entry into the nucleus, increasing the chance of genomic integration upon antibiotic selection.
1. Seed 2-3 3 10 6 C17.2 cells per 15 cm cell culture dish (1 for each plasmid nucleofection) and incubate at 37 C/ 5% CO 2 overnight resulting in 3-4 3 10 6 cells/ 80% confluency. 2. Remove the culture medium from the cells and wash cells once with Dulbecco's phosphate-buffered saline (DPBS), using at least the same volume of PBS as culture medium. 3. Proceed with harvesting by incubating cells at 37 C with trypsin/EDTA for 3 min or until all cells are detached from the culture dish surface. 4. Add complete medium to the dish, and pipette up and down until you get a homogenous singlecell suspension. 5. Take 10 mL from the single-cell suspension and determine the cell density with a hemocytometer. 6. Centrifuge 1 3 10 6 cells/condition at 200 g for 10 min at room temperature (RT). During this time, prepare nucleofection materials: a. Add Supplement solution to the Nucleofectorä Solution (supplied in the kit). b. Start 4D-Nucleofectorä System (Lonza) and upload DN-100 experimental parameter file as for C17.2 cells, SG transfection solution and Nucleofectorä program DN100 showed good results (refer to this protocol https://bioscience.lonza.com/lonza_bs/CH/en/download/ product/asset/21623, however note that the program is DN100 instead of that mentioned in the protocol, follow the procedure for 100 mL Nucleocuvetteä). c. Fill 10 mL complete culture medium in each 10 cm cell culture plate (one plate per condition) and pre-incubate/equilibrate plates in a humidified 37 C/5% CO2 incubator. d. Pre-warm an aliquot of culture medium to 37 C. Prepare plasmid DNA. 7. After 10 min of centrifugation, remove the supernatant and resuspend the cell pellet carefully in RT 4DNucleofectorä Solution. 8. Add 5 mg of plasmid DNA to this suspension (max. 10% of final sample volume) and transfer to a Nucleocuvetteä Vessel. 9. Repeat this procedure for the rest of Nucleocuvetteä Vessels as required.
Note: As prolonged incubation of cells in Nucleofectorä Solution may lead to reduced transfection efficiency and viability it is important to work as quickly as possible. Avoid air bubbles while pipetting.
CRITICAL: Make sure to have <85% cell confluency when collecting cells for nucleofection. Cell confluency >85% will result in reduced transfection efficiencies.
Note: C17.2 cells tend to grow faster at higher passage number, and eventually tend to differentiate into elongated cells that stop dividing. Hence, it is advisable to use cells with low passage number (<15) to generate a stable cell line by means of nucleofection.
10. Tap the Nucleocuvetteä Vessels gently to cover the sample at the bottom of the cuvette. 11. Close the lid and place the cuvette into the retainer of the 4D-Nucleofectorä X Unit in proper orientation. 12. Press Start on the display of the 4D-Nucleofectorä Core Unit (for details, please refer to the device manual https://bioscience.lonza.com/lonza_bs/NL/en/document/31856). 13. Remove the cuvette from the retainer after completing the run. Note: In our hands, it was 50% with 40%-60% cell viability.
18. Replace the complete medium with the one supplemented with 3 mg/mL puromycin.
Note: Generally, 1 mg/mL puromycin is sufficient for polyclonal cell line preparation. However, to select cells with high expression of recombinant protein, a higher puromycin concentration is recommended).
19. Over the next few days non-transfected cells will die. To remove dead cells and cell debris, refresh the medium with fresh medium with puromycin every other day. 20. After a couple of weeks, only cells that have incorporated the exogenous DNA into their genome will have survived the antibiotic selective pressure. A representative fluorescence micrograph of cells expressing XP-GFP-DNAJB6 is shown in Figure 3.  They should grow apart from each other as this is important for picking single colonies later on. 27. There should be at least 100 cells per colony. Keep checking the size of the colonies every now and then. Generally, colonies will be ready for picking 2 weeks post-seeding. 28. In the meantime, cut out Whatman paper (number 1) discs with a punching machine and autoclave them. 29. At the day of colony picking, seal the dish with parafilm. Clean the microscope stage with 70% ethanol and screen the dish for colonies with GFP fluorescence. Encircle the colonies (> 12 clones) with highest fluorescence on the bottom of the dish using a marker pen. 30. Remove the parafilm and place the dish back in the incubator. Warm up cell culture medium, trypsin and HBSS. Take 5 mL trypsin in a small dish. Put 12-15 autoclaved paper discs in the lid of the dish. 31. Distribute 500 mL complete medium per well in a 24 well plate. Label the wells with numbers. 32. Take the dish with colonies out of the incubator and place it in the laminar flow. Wash with HBSS twice. Flame the tweezers twice and let them cool down. 33. Meanwhile, remove HBSS from the dish. From this time onwards, one has to work fast in order to prevent cells from drying. Take a paper disc, dip it in trypsin/EDTA and put on the encircled colony. Repeat this for all the colonies. Keep the dish in the incubator for 3 min. 34. Pick the paper discs one by one and put one disc per well in the 24 well plate. 35. After transfer of all discs to the 24 well plate, pipette up and down medium in every well with a separate tip with a 1 mL micropipette to rinse the discs with the medium. 36. Place the 24 well plate in the incubator. 37. Keep checking this plate daily. It will take 4-5 days before cells are detected in the wells. Once the wells are around 70% confluent, discard the Whatman paper discs, transfer cells from one well in 24 well plate to one well in 6 well plate. 38. Select clones that show the maximum amount of GFP and GFP-DNAJB6 expression at the expected localization of XPackä tagged proteins, i.e., at the plasma membrane and endosomal structures.
Note: These clones are your parental cell lines for XP-GFP and XP-GFP-DNAJB6 loaded extracellular vesicle production, respectively.
39. Transfer each of the selected clones to a separate 10 cm dish, and let cells grow until 80% confluence. Expand each 10 cm dish to 3 10 cm dishes. Freeze cells from 2 dishes in 5 cryogenic vials. Use the remaining 10 cm dish for downstream processing.
CRITICAL: Expression of XPackä tagged GFP fusion proteins produces a cellular distribution pattern similar to sEV marker proteins, e.g., the tetraspanin CD9. Specifically, some of the GFP signal is localized on the plasma membrane and most is localized at multivesicular endosomes. 1

Timing: 30 min
This section describes the preparation of cleaning coverslips for microscopy ( Figure 4).
We use 13 mm glass coverslips (VWR) to seed cells on in 24 well plates. For quick and effective cleaning, we advise the procedure below which involves dipping the coverslips in 100% ethanol and flaming them to remove dirt and for sterilization.  CRITICAL: We recommend always making fresh sEV isolates for functional assays, including protein aggregation studies. You may store them overnight at 4 C, and use them the next day. However, do not keep them at 4 C for an extended period of time.
We have observed that sEVs tend to aggregate over time. Storage of sEVs at -20 C is not recommended either because of the potential loss of functional activities. 7 For nonfunctional assays such as sEV marker characterization with western blotting, you may freeze the samples in loading buffer and store at -80 C. Please refer to 57-69 for further details.

OPEN ACCESS
57. 48 h later, collect the medium (180 mL) in 5 UC tubes (36 mL/tube) and isolate extracellular vesicles by sequential centrifugation as follows.
58. Add an equal amount of sterile water in 1 UC tube which will be for balancing. Weigh the tubes to balance and proceed with centrifugationa. Remove dead cells from the supernatant by centrifugation at 500 g for 10 min at 4 C. Carefully transfer the supernatant to a fresh falcon tube by pipetting without touching the pellet. The procedure can be performed at RT. b. Next, centrifuge the supernatant at 2,000 g for 10 min at 4 C to remove cellular debris (Beckman Coulter, Allegra X-15R). Carefully transfer supernatants into a UC tube (38. 67. Add 300-500 mL of antibody solution (prepared in odyssey blocking buffer, 1:1000) on the parafilm. 68. Take out the blot from the blocking solution, cut a small piece off right corner to mark the protein side, and place the blot with the protein side facing the antibody solution i.e., facing downward. 69. Place one small wet tissue paper along the edges of the petri dish. Make sure that the tissue paper is not touching the blot, to prevent drying of the blot by capillary action. 70. Close the dish, and incubate the blots overnight at 4 C. 71. Next day, wash the blots with 0.1% PBS-Tween20 4 3 5 min. 72. Incubate the blots with secondary antibody solution (1:5000 in blocking buffer) for 1 h at RT, following the same procedure as for primary antibody incubation. 73. Wash with 0.1% PBS-Tween20 4 3 5 min. 74. Take out the blot from the petri dish and acquire images with an Odysseyâ Infrared Imaging System (Li-COR) following manufacturer's protocol (https://www.licor.com/bio/support/contents/ applications/western-blots/fluorescent-western-blot-detection-protocol.html?Highlight=on-cell western assay).

Timing: 4 days
This protocol describes the procedure to make an in vitro HD cell model to assess the potency of DNAJB6-containing sEVs using fluorescence microscopy.
CRITICAL: As this is a functional assay, it is important that you use freshly isolated sEVs. We recommend seeding cells for the inclusion body assay on the day you prepare the western blot samples to check for the presence of DNAJB6 and EV markers. When their presence is confirmed, use the sEV isolates within 1-2 days.
CRITICAL: It takes >8 h post-transfection for protein expression. As DNAJB6 acts upstream of aggregation i.e., it prevents aggregate formation but does not remove existing aggregates, it is important that you add sEV solution right after the jetPEI incubation for best results. For cargo proteins acting downstream of protein aggregation, sEVs can be added later after transfection (e.g., 24 h post-transfection). Note: As the inclusion bodies show very high intensity signal, it is advisable to start with low intensity of excitation. See Figure 5 for reference.

Analysis of inclusion body numbers -
a. Count the number of nuclei using ImageJ Plugins > Analyze > Cell counter. Make sure that you only count cells that have been transfected. b. Count the number of inclusion bodies using the same plugin. c. Document all the numbers in an excel file. d. Get a ratio of the number of cells with inclusion bodies to the total number of transfected cells. 87. Repeat the experiment 3 times. Draw a bar graph using Graphpad PRISM or any other software for analsysis.

Timing: 4 days
This protocol explains the filter trap assay, which separates aggregated (insoluble) proteins from soluble proteins. Shortly, cell lysates ( Figure 6) are passed through the cellulose acetate membrane which traps aggregates >200 nm, separating them from soluble proteins (Figure 7). The trapped proteins can be detected, which enables semiquantitative measurements for protein aggregation.    Note: It is important that you make these samples from freshly made isolates. However, once these are boiled with the loading buffer, they can be stored at -80 C until further use. However, avoid freeze-thaw cycles until use.
109. For western blotting, take 30 mg of total protein of each sample from step 99 and follow steps 61-74 using anti-GFP antibody with the following modifications. a. As the protein aggregates are too big to travel through running gel, they are trapped in the stacking gel. Hence, it is important to use the entire gel for transfer onto the PVDF membrane. b. The stacking gel easily breaks during transfer, hence lift and place the gel gently on the membrane. Rearrange the well walls into the original position if necessary. 110. Analysis of the western blot to quantify protein aggregation using ImageJ.
a. Draw a rectangle around the bands in ImageJ in such a way that you can have a rectangle of the same size for all bands. b. Using Analyze > Measure, measure the band intensities of the protein aggregation in each sample. c. Calculate the extent of protein aggregation as a percentage with respect to control.

EXPECTED OUTCOMES
The clinical translation of therapeutics for brain diseases, including Huntington's disease, is hindered by the generally poor accumulation of therapeutics in the brain upon their systemic administration. Drug delivery vehicles are being developed to facilitate drug delivery at the target site. sEVs have emerged as promising biological delivery vehicles for various biomolecules, including proteins. 8 In this protocol, we have described the procedure of loading sEVs with the potent anti-aggregation chaperone protein DNAJB6, and demonstrate their use in reducing aggregation in an in vitro cellular model for HD.
There are two main outcomes of this protocol -high loading of DNAJB6 in sEVs, and reduction in the amount of protein aggregation in polyQ expressing cells.
High loading of DNAJB6 in sEVs It is difficult to determine the exact amount of loaded DNAJB6 protein in each EV without performing further specialized biochemical experiments. If 10 mg of EVs can give a detectable signal of DNAJB6 in western blotting, the DNAJB6 expression levels would be sufficient to produce detectable decrease in protein aggregation in cells in the assays stated in this article. Hence, make sure to pick colonies that show maximum DNAJB6-GFP signal to maximize the probability of getting high loading in sEVs.
Reduction in the amount of protein aggregation in polyQ expressing cells In our hands the DNAJB6-loaded sEVs generated 30% decrease in protein aggregation in HD cellular model in both fluorescence microscopy and WB analyses.

LIMITATIONS
The most problematic limitation, is the high volumes of conditioned media required to get sufficient amounts of sEVs for experiments. Although C17.2 cells produce large amounts of sEVs, bioreactors may be considered for higher and less labor-intensive production of sEVs.
In addition, due to (high) protein overexpression, overall cellular function may be impacted. As a consequence, it is possible that the EVs contain not only the protein of interest but also other components that can be either beneficial or non-beneficial to the process under study. Potential solution C17.2 cells are known to resist exogenous expression. But it is important to have high amounts of DNAJB6 in cells in order to maximize the sEV loading. An option is to use high(er) amounts of plasmid DNA during nucleofection. You may also increase the antibiotic concentration to 5 mg/mL in order to select only high DNAJB6 expressing cells.

Potential solution
During sEV isolation procedure, the main loss takes place during washing of the sEV samples. One way to avoid this loss is to use less washing steps during ultracentrifugation or use smaller ultracentrifugation tubes. Another reason for a low sEV yield is the incomplete recovery of sEV pellets due to long handling times, especially when pellets are left in buffers for a longer period of time before resuspending. Hence, try to avoid long handling times, or leave the pellets in wash buffers instead of media supernatants to avoid loss during transfer.

Potential solution
We have often experienced variability in the sEV field. We have found that the major cause of this is variability in cell seeding and higher passage number. Use of different ultracentrifugation equipment within experiments also results in variability. Hence, try to attain similar and optimal cell density before harvest and use cells with passage number <40. Try to use the same equipment for all the isolates.

Problem 4
Variable suppression levels in protein aggregation assay (step 108).

Potential solution
Not often, but a few times, we have experienced some variability in the suppression levels of protein aggregation among different biological samples. The main cause of this is different quality of sEV samples. It is important to characterize the sEV isolate every time to confirm the levels of DNAJB6 and EV markers. If they attain the values similar to all other experiments, then only proceed to functional assays.

Lead contact
Requests for resources and reagents should be directed to the lead contact, Inge Zuhorn (i.zuhorn@ umcg.nl).

Materials availability
Plasmids and cell lines are available upon reasonable request.

Data and code availability
This study did not generate/analyze datasets/code.